Single-Molecule Measurements of the Binding between SmallMolecules and DNA AptamersPhilip M. Yangyuoru,‡ Soma Dhakal,‡ Zhongbo Yu,‡ Deepak Koirala,‡ Simon M. Mwongela,†

and Hanbin Mao*,‡

†School of Science & Technology, Georgia Gwinnett College, Lawrenceville, Georgia 30043, United States‡Department of Chemistry and Biochemistry, Kent State University, Kent, Ohio 44242, United States

*S Supporting Information

ABSTRACT: Aptamers that bind small molecules can serve asbasic biosensing platforms. Evaluation of the binding constantbetween an aptamer and a small molecule helps to determinethe effectiveness of the aptamer-based sensors. Bindingconstants are often measured by a series of experiments withvarying ligand or aptamer concentrations. Such experiments aretime-consuming, material nonprudent, and prone to lowreproducibility. Here, we use laser tweezers to determine thedissociation constant for aptamer−ligand interactions at thesingle-molecule level from only one ligand concentration.Using an adenosine 5′-triphosphate disodium salt (ATP) binding aptamer as an example, we have observed that the mechanicalstabilities of aptamers bound with ATP are higher than those without a ligand. Comparison of the change in free energy ofunfolding (ΔGunfold) between these two aptamers yields a ΔG of 33 ± 4 kJ/mol for the binding. By applying a Hess-like cycle atroom temperature, we obtained a dissociation constant (Kd) of 2.0 ± 0.2 μM, a value consistent with the Kd obtained from ourequilibrated capillary electrophoresis (CE) (2.4 ± 0.4 μM) and close to that determined by affinity chromatography in theliterature (6 ± 3 μM). We anticipate that our laser tweezers and CE methodologies may be used to more conveniently evaluatethe binding between receptors and ligands and also serve as analytical tools for force-based biosensing.

The interaction between nucleic acids and small moleculesis continuously generating significant interest due to their

widespread use in analytical and bioanalytical applications.Specific nucleic acids such as aptamers have been shown tobind not only to nucleic acids but also to non nucleic acidtargets such as small molecules,1,2 proteins,3 and cells.4

Aptamers are single-stranded ligand-binding nucleic acidswith affinity and selectivity comparable to those of antibod-ies.5,6 Compared to antibodies, DNA aptamers are easier toobtain, more stable, and more resistant to biodegradation. Assuch, they are suitable for demanding conditions such asextreme pH and temperatures. These properties render themideal candidates for sensors used in medical diagnostics andenvironmental monitoring.7

The binding affinity between an aptamer and a ligand is acritical factor to reflect the sensitivity of aptamer-basedbiosensors.8 Conventionally, the binding is measured bytechniques such as electrochemical detection,9,10 fluores-cence,11 colorimetric12 or surface plasma resonance (SPR)based detection,13 and capillary electrophoresis (CE) separa-tion,14−16 among others.17 At the single-molecular level, atomicforce microscopy (AFM) has been recently used to determinethe adenosine 5′-triphosphate disodium salt (ATP) binding toan aptamer made of two independently split DNA strands.18

Single-molecule fluorescence resonance energy transfer(FRET) has also been explored to evaluate the binding of a

DNA aptamer with its binding target, vascular endothelialgrowth factor (VEGF).19

However, a majority of these methods require a series ofexperiments with varying ligand or aptamer concentrations todetermine the dissociation constant. Apart from being tediousand time-consuming, such experiments are prone to adecreased signal-to-noise ratio due to run-to-run variations.Another disadvantage is that they require a substantial amountof materials. This becomes problematic especially in drugscreening processes where the amount of a sample is oftenlimited. As an alternative, here, we employ a laser tweezersinstrument to determine the dissociation constant betweenATP and a DNA aptamer by using only one concentration ofATP.Developed in the 1980s,20 laser tweezers have been used

extensively to probe the mechanical stabilities not only forDNA21,22 and RNA23−26 but also for proteins27 at the single-molecule level. Recently, we have extended this method toinvestigate the interaction between a DNA G-quadruplex andits ligand,28 as well as to serve as a force-based biosensingplatform.29 Here, we use this technique to scrutinize theinteraction between aptamers and ligands. The binding of ATP

Received: February 13, 2012Accepted: May 22, 2012Published: May 23, 2012

Article

pubs.acs.org/ac

© 2012 American Chemical Society 5298 dx.doi.org/10.1021/ac300427d | Anal. Chem. 2012, 84, 5298−5303

to the aptamer is expected to increase the mechanical stabilityof the aptamer. This increment is converted to the change inthe free energy of binding (ΔGbinding) by a Hess-like process.The ΔGbinding is then used to obtain the dissociation constant(Kd) of the complex, which is validated by our CEmeasurement. Such a force-based strategy is less prone tobackground noise29 often seen in fluorescence-based methods.Our results demonstrate a strong correlation between ensembleaverage assays (CE) and single-molecule approaches (lasertweezers) on the binding between aptamers and smallmolecules.

■ MATERIALS AND METHODS

Materials. ATP was purchased from USB Corp., Cleveland,OH. N-[Tris(hydroxymethyl)methyl]glycine (Tricine) waspurchased from Fisher Scientific. Trizma base (minimum99.9% titration) was purchased from Sigma. Poly-(vinylpyrrolidone) (PVP) was obtained from Acros Organics.Nanopure water was purified by Millipore Synergy UV with a185 nm UV lamp. The ATP aptamer and a scrambled sequencefor control experiments were labeled with 6-carboxyfluorescein(FAM) to form 5′-[FAM]TAC CTG GGG GAG TAT TGCGGA GGA AGG TT-3′ and 5′-[FAM]TAG TGC GCG TGTGTG AGC AGA GTG AGA GT-3′, respectively. They werepurified using HPLC by the manufacturer (Integrated DNATechnologies (IDT), Coralville, IA). Other DNA fragmentswere purchased from IDT and purified with polyacrylamide gelelectrophoresis (PAGE).Synthesis of the DNA Construct for Single-Molecule

Studies. Preparation of the DNA construct for single-moleculemechanical unfolding studies was carried out following aprevious protocol.30 Overall, an ATP DNA aptamer fragmentwas sandwiched between two double-stranded DNA (dsDNA)handles (2690 and 2028 bp). The 2690 bp handle wasconstructed from a pEGFP plasmid (Clontech, Mountain View,CA) by SacI (New England Biolabs (NEB), Ipswich, MA) andEagI (NEB) digestions. It was gel purified and labeled withdigoxigenin (Dig) at the 3′ end (SacI end) using 18 μM Dig-dUTP (Roche, Indianapolis, IN) and terminal transferase(Fermentas, Glen Burnie, MD). To ensure complete Diglabeling, excess Dig-dUTP was used (Dig-dUTP:2690 bp DNA= 150:1). The Dig-labeled 2690 fragment was purified byethanol precipitation. To make the 2028 bp handle, a pBR322plasmid (NEB) was first amplified by polymerase chain reaction(PCR) using a 5′-labeled biotinylated primer, 5′-GCA TTAGGA AGC AGC CCA GTA GTA GG-3′ (IDT). The PCRproduct was purified by a kit (Qiagen, Germantown, MD) anddigested overnight with XbaI restriction enzyme (NEB).The 83-mer DNA fragment sandwiched by the 2028 and

2690 bp handles contains a single-stranded ATP aptamersequence in the middle, 5′-ACC TGG GGG AGT ATT GCGGAG GAA GGT. This sequence is flanked by two dsDNAfragments that end with 5′-GGC CGA CGC GCT GGG CTACGT CTT GCT GGC and 5′-CGC ATC TGT GCG GTATTT CAC ACC GT sticking ends, to which the 2028 and 2690bp dsDNA handles were ligated using T4 DNA ligase (NEB)and purified using a gel extraction kit (Qiagen). The finalconstruct was ethanol precipitated, dissolved in water, andstored at −20 °C. The DNA construct with a scrambledsequence, 5′-AGT GCG CGT GTG TGA GCA GAG TGAGAG, in place of the ATP aptamer, was prepared by the sameprocedure.

Single-Molecule Experiments. A detailed description ofthe laser tweezers instrument used for the single-moleculeexperiment has been reported elsewhere.31 To start the single-molecular experiment at room temperature, the appropriatelydiluted DNA construct prepared above (0.50 ng/μL) wasincubated with anti-Dig-coated polystyrene beads (diameter2.10 μm, Spherotech, Lake Forest, IL) for 1 h at roomtemperature. This allowed the DNA construct to bind to thebeads through the Dig−anti-Dig linkage. The incubated samplewas diluted to 800 μL in a 10 mM phosphate buffer (pH 7.4)with 100 mM KCl. Streptavidin-coated beads (diameter 0.97μm, Bangs Laboratory) were dispersed into the same workingbuffer. The streptavidin-coated beads and the anti-Dig-coatedbeads linked with the DNA construct were injected separatelyinto a reaction chamber through different channels. These twotypes of beads were captured separately by the two laser traps.The two laser-trapped beads were moved to touch against eachother to allow the free end of a DNA molecule on the anti-Dig-coated bead to tether to the streptavidin-coated bead via thestreptavidin−biotin linkage. The anti-Dig-coated bead, con-trolled by the Nano-MTA positioned in the laser tweezersinstrument, was then moved away from the streptavidin-coatedbead with a loading speed of ∼5.5 pN/s. A sudden drop in thetension was observed whenever there was an unfolding of asecondary structure. Single tethers were confirmed by observingthe single breakage of the tether during stretching or the 65 pNplateau in force−extension (F−X) curves.32 The force−extension curve for each tether was recorded in a Labviewprogram, and data treatment was performed using programs inMatlab and Igor software. Only F−X curves from single tetherswere considered for data analysis. Two data points flanking therupture event in the F−X curves were used to calculate thechange in contour length (ΔL) during the rupture of a foldedstructure according to the following equation:22

= − +⎛⎝⎜

⎞⎠⎟

xL

k TFP

FS

112

B1/2

(1)

where x is the end-to-end distance, L is the contour length, T isabsolute temperature (298 K), F is the force, kB is theBoltzmann constant, P = 51.95 nm33 is the persistent length,and S = 1226 pN33 is the elastic strength modulus.The bias of these free energy estimations was calculated

according the described method.34 The dissipated work, 12 and15 kBT for unfolding at 0 and 100 μM ATP, respectively, waswell within the range (∼20 kBT) for effective bias calculations.34The formation probability of a folded species is calculated by

taking the ratio of F−X curves that contain rupture eventsversus the total number of tethered F−X curves.

Capillary Electrophoresis Experiments. Aliquots of ATP(0−60 μM) were added to the ATP aptamer (0.5 nM) in a 10mM phosphate buffer (pH 7.4) with 100 mM KCl. The mixturewas heated to 97 °C for 5 min followed by slow cooling toroom temperature for 4 h in a Bio-Rad DNAEngine Peltierthermal cycler. Capillary electrophoresis was performed on aBeckman ProteomeLab PA800 CE system (Fullerton, CA).The machine is equipped with a laser-induced fluorescence(LIF) detector using an air-cooled 3.5 mW argon laser(Beckman Instruments), with excitation at 488 nm andemission at 520 nm, and is installed with 32 Karat 8.0 software(Beckman Instruments) for instrument control and dataanalysis. Separations were performed on 30 cm fused-silicacapillaries (Polymicro Technologies, Phoenix, AZ) with 50 μm

Analytical Chemistry Article

dx.doi.org/10.1021/ac300427d | Anal. Chem. 2012, 84, 5298−53035299

i.d. and 375 μm o.d. (20 cm effective length). New capillarieswere preconditioned by successively flushing with 1.0 M NaOHfor 30 min and rinsed with distilled deionized H2O for 20 minat 20 psi. Samples were injected hydrodynamically at 1.0 psi for10 s. The separation buffer was made of 25 mM Tris, 25 mMTricine, and 0.5% PVP. During the separation the capillarycolumn was thermostated at 25 °C, and an electric fieldstrength of 500 V/cm was applied in the reverse polarity mode(anode at the detector).

■ RESULTSEnhancement of the Mechanical Stability of an

Aptamer by Small-Molecule Binding. We first investigatedthe mechanical stability of the ATP aptamer itself. Aftersandwiching the aptamer fragment between two dsDNAhandles, the DNA construct was tethered between the twolaser-trapped beads via biotin−streptavidin and Dig−anti-Digantibody linkages (Figure 1A). The tension in the DNA tether

was gradually increased by moving one of the trapped beadsaway from the other until a sudden drop in force was observedin the F−X curves (Figure 1C inset). This drop in force isassociated with the unfolding of a secondary structure, mostlikely the aptamer, in the DNA construct. The change incontour length (ΔL; see the Materials and Methods for thecalculation) was determined to be 9.2 ± 0.1 nm, a valueconsistent with the contour length of the 27-mer ATPaptamer.30,35 In a control with a scramble sequence (see theMaterials and Methods) that does not assume any stableconformations, no folded structure was observed (data notshown), which strongly suggests that the observed events aredue to the ruptures of folded ATP aptamers. The unfoldingforce at which the rupture event occurred showed a mostprobable value of 21 pN without the ATP ligand (Figure 2A).The probability of observing these rupture events from alltethered DNA molecules is 24% (see the Materials andMethods for the calculation), suggesting a significant fraction ofthe aptamers is not folded.To investigate the binding of a small molecule, the aptamer

construct was incubated with 2.0 μM ATP. The rupture forcehistogram for this treatment revealed two populations with

rupture forces centered at 21 and 38 pN (Figure 2B). On thebasis of the similar rupture force, we assigned the 21 pNpopulation as the free aptamer. After the aptamer binds to ATP,it is expected that the mechanical stability of the aptamer willincrease.28 Therefore, the 38 pN species was assigned as theATP-bound aptamer (Figure 2B). To confirm this assignment,we increased the ATP concentration to 100 μM with theexpectation that the population of the ATP-bound aptamershould increase. Indeed, this was observed in the unfoldingforce histogram in Figure 2C. The increase in the mechanicalstability of the aptamer upon ligand binding can be exploited tofurther develop force-based biosensing recently developed inour laboratory.29

Dissociation Constant Determination at Only OneConcentration of a Ligand. The dissociation constant (Kd)between the ATP and the aptamer can be calculated by theexpression ln(Kd) = −ΔGbinding/RT, where ΔGbinding is thechange in free energy of binding, a state function that isindependent of pathways. To obtain the ΔGbinding, we designeda Hess-like cycle28 in which the binding of the ligand to theaptamer is rerouted to the unfolding of a free aptamer in thepresence of an unbound ligand. This is followed by theformation of the ATP-bound complex from the mechanicallyunfolded aptamer (Figures 3 and 4). The ΔGbinding is thenestimated from the difference in the change in free energy ofunfolding (ΔGunfold) for the latter two processes. Each ΔGunfoldis obtained using Jarzynski’s equality equation36

∑Δ = − −−

⎡⎣⎢

⎤⎦⎥G k T

NW

k Tln

1exp

i

Ni

unfold B1 B (2)

where kB is the Boltzmann constant, T is absolute temperature,N is the number of observations, and W is the nonequilibriumwork done to unfold the structure(s), which is equivalent to thehysteresis area between stretching and relaxing F−X curves (seeFigure 1C as an example). Histograms for the work revealedthat the average work done to unfold the free aptamer (Figure3A) is less than that for the ATP-bound aptamer (Figure 3B),which confirms that the mechanical stability of the aptamer is

Figure 1. (A) Schematic of the experimental setup. (B) ATP structure.(C) Typical force−extension curve. The inset shows a blowup of theunfolding event denoted by the dotted circle.

Figure 2. Unfolding force histograms of the ATP aptamer in a 10 mMphosphate buffer (pH 7.4) with ATP concentrations of (A) 0 μM, (B)2.0 μM, and (C) 100.0 μM. Curves depict Gaussian fittings.

Analytical Chemistry Article

dx.doi.org/10.1021/ac300427d | Anal. Chem. 2012, 84, 5298−53035300

enhanced after ligand binding. The Jarzynski calculation yieldeda ΔGfree‑unfold of 29 ± 3 kJ/mol (bias estimation 7 kJ/mol) and aΔGbound‑unfold of 62 ± 3 kJ/mol (bias estimation 8 kJ/mol).With these, ΔGbinding was calculated by the followingexpression:

Δ = Δ − Δ‐ ‐G G Gbinding free unfold bound unfold (3)

This treatment yielded a ΔGbinding of −33 ± 4 kJ/mol, whichled to a Kd of 2.0 ± 0.2 μM by the equation ΔGbinding = −RT lnKd.Determination of the Dissociation Constant by CE. To

validate these single-molecule results, we set out to compare theKd value with an ensemble measurement. It is known thatbinding affinity can be determined by capillary electro-phoresis.37,38 Therefore, the ATP aptamer was labeled with 6-carboxyfluorescein at the 5′ end and analyzed using CE-LIF.Depicted in Figure 5 are two populations (∼4.6 and ∼4.2

min retention times) in the electropherogram of the 0.5 nMATP aptamer with increasing ATP concentrations (0−60 μM).Since a significant fraction of DNA is not folded as revealed bylaser tweezers experiments (see above), we ascribe the largerpopulation (the 4.2 min peak) as unfolded aptamer and the 4.6min population as the folded aptamer bound with or withoutATP. The difference in the elution time reflects the variation inthe size and charge between folded and unfolded ATP

aptamers, which is likely due to different interactions withpotassium cations.39,40 This assignment was confirmed as thefolded aptamer species (the 4.6 min peak with free and ATP-bound aptamers) increase in population relative to the unfoldedones when the ATP concentration increases (Figure 5, 0−60μM ATP traces). As a control, a scrambled sequence (see theMaterials and Methods) that does not form any structures didnot show this trend (see Figure S-1 in the SupportingInformation). The fact that neither unfolded nor foldedaptamers had tailing or fronting in the electropherogramsuggests that the conversion between these two species is slowwithin the time scale of electrophoresis. In addition, theinseparable ATP−aptamer complex and the free folded aptamerhelp to maintain the equilibrium between the free and the ATP-bound aptamers during the electrophoresis. These two factsdirectly contrast with the nonequilibrated conditions exploitedby kinetic CE for measurement of dissociation constants.41,42

These equilibrated conditions were used to determine thedissociation constant for the ATP−aptamer complex. First, wemeasured the amount of folded aptamer without bound ATPfrom the 4.6 min peak in the electropherogram at 0 μM ATP.By comparing the folded aptamer with the unfolded aptamerthat appeared at the 4.2 min peak, we obtained a foldingequilibrium constant of 0.30 (Kfold = [folded]/[unfolded]).This Kfold value is very close to that estimated from theformation probability of folded species (24%) observed in thesingle-molecule studies ([probability of folded]/[probability ofunfolded] = 24%/76% = 0.32). With this Kfold value, we thendetermine the fraction of the free, folded aptamer in the 4.6min peak by comparison with the unfolded aptamer population(the 4.2 min peak) for all other ATP concentrations (Figure 5,0−60 μM ATP traces). On the basis of these fractions, aLangmuir isotherm28 (Figure 5B) was constructed to calculatethe Kd of the ATP−aptamer complex. The resultant Kd (2.4 ±0.4 μM) matches very well with the single-molecule Kdmeasurement obtained above. It is also comparable withprevious measurements based on affinity chromatography.43

Figure 3. Histograms of the work done to unfold the free aptamer (A)and ATP-bound aptamer (B). The work is calculated at 100 μM ATP.Solid curves depict Gaussian fittings. Dotted arrows show the workequivalent to the change in free energy of unfolding (ΔGunfold).

Figure 4. Hess-like cycle of the aptamer−ATP binding process usedfor the calculation of the ΔGbinding and the determination of thedissociation constant.

Figure 5. (A) Electropherograms for 0.5 nM aptamer in the presenceof varying ATP concentrations (0−60 μM) in a 10 mM phosphatebuffer (pH 7.4) containing 100 mM KCl at 25 °C. The left peak (4.2min) represents the unfolded aptamer, while the right one (4.6 min)indicates folded aptamer species (free and ATP-bound). (B) Fractionof the ATP-bound aptamer with ATP concentration. The solid curve isa Langmuir isotherm fitting to obtain the dissociation constant, Kd.

Analytical Chemistry Article

dx.doi.org/10.1021/ac300427d | Anal. Chem. 2012, 84, 5298−53035301

■ DISCUSSION

Kd Determination by Capillary Electrophoresis. Pio-neered by Krylov, kinetic CE has been developed in the past 10years to obtain the dissociation constants from nonequilibriumconditions created during capillary electrophoresis.41 Thismethod requires different electrophoretic mobilities betweenthe ligand, the receptor, and the ligand−receptor complex. Inaddition, the binding or unbinding should be fast enough toallow the interaction to occur within the time scale of the CEseparation. Here, due to the inseparable species of bound andfree aptamers, and the slow folding and unfolding of theaptamer (see above), our method allows the determination ofthe binding affinity in a manner different from that of kineticCE approaches. This equilibrated CE method is also differentfrom the previous CE approaches that utilize the differentmobilities between the free and bound species to obtain thebinding affinity.14 Our CE method provides a Kd value close tothat from affinity chromatography (6 ± 3 μM),43 validating ourequilibrated CE approach for the affinity measurement.Kd Determination by Laser Tweezers. Single-molecule

studies can reveal detailed mechanisms of the interactionbetween ligands and receptors.44 However, not many single-molecule approaches are capable of retrieving thermodynamicvariables, such as Kd, which is best obtained at equilibriumestablished by millions of molecules. Being highly sensitive,single-molecule techniques are better suited to investigatekinetics processes. Jarzynksi’s approach45 to retrieve thermody-namic information from nonequilibrated processes greatlyexpands the capability of single-molecule techniques. Ourmethods described above fully utilize this innovative discovery.By repeatedly unfolding and refolding individual structures witha laser tweezers instrument, the change in free energy ofunfolding can be obtained from nonequilibrium conditions.The single-molecule setting reduces the number of unfoldingand refolding cycles that are required to retrieve thethermodynamic information as the thermal bath from theenvironment becomes significant at the single-molecular level.The validity of such an approach has been well establishedpreviously.45

The fact that the binding energy, ΔGbinding, which is used forthe dissociation constant calculation, can be obtained in thepresence of a ligand with a desired concentration enables ourmethod to determine the binding affinity from only oneconcentration of the ligand (Figure 3). Previously, methodshave been demonstrated to measure the affinity from a ligandwith a fixed concentration.46,47 However, all these approachesrequire a certain range of workable concentration to ensureaccurate determination of a dissociation constant. In our case,this limitation is not necessary. In practice, we can choose aligand concentration high enough to populate ligand-boundspecies, from which the ΔGunfold is determined (in this research,we obtained ΔGunfold from the 100 μM ATP concentration).After correction for the ΔGunfold for an aptamer without abound ligand, ΔGbinding is obtained (Figure 4). Therefore, ourmethod can be conveniently used to evaluate the dissociationconstant without prior knowledge.This single-molecule approach provides a general platform to

measure the binding affinity of a receptor−ligand systembeyond the aptamer−ligand complex described here. Toinvestigate protein−ligand interaction, a protein can betethered between two optically trapped particles. Themechanical stability of the protein can then be evaluated with

or without a ligand to determine the dissociation constant usingthe Hess-like cycle (Figure 4). Recently, several proteins48,49

have been investigated for their mechanical stabilities by lasertweezers using a similar molecular-construct strategy describedhere. Compared to the fluorescence-based evaluations that areprone to background noises, the force-based detection hasreduced noise.29 In addition, the positions from which areceptor is attached to handles can be well controlled to avoidthe labeling impact on the binding pocket, which is commonfor fluorescence-based methods. Finally, the aptamer-contain-ing single-molecule system serves as a natural platform forforce-based sensing. Previously, we have described detection ofsingle-nucleotide polymorphism (SNP) using this strategy.29 Byexploiting the versatility of aptamers, different small moleculescan be detected on the basis of the increased mechanicalstability of a ligand-bound aptamer.

■ SUMMARYIn summary, we have used two complementary methods, anensemble-average-based CE and a single-molecule-basedmechanical unfolding, to investigate the binding of ATP toan ATP aptamer. The aptamer−ligand interaction contributesto the increased mechanical stability of the complex, whichleads to the determination of the dissociation constant by aHess-like cycle. This dissociation constant is consistent withthat determined by our equilibrated CE analysis and close tothat measured in the literature.50 We anticipate the methodsdeveloped here not only are useful for the determination of thebinding affinity of general receptor−ligand systems in aconvenient manner but also can be used to develop force-based biosensors.

■ ASSOCIATED CONTENT*S Supporting InformationAdditional information as noted in text. This material isavailable free of charge via the Internet at http://pubs.acs.org.

■ AUTHOR INFORMATIONCorresponding Author*E-mail: E-mail:hmao@kent.edu. Phone: (330) 672-9380.

NotesThe authors declare no competing financial interest.

■ ACKNOWLEDGMENTSWe thank the National Science Foundation (Grant CHE-1026532) for financial support of this work.

■ REFERENCES(1) Ellington, A. D.; Szostak, J. W. Nature 1992, 355, 850−852.(2) Drabovich, A. P. B., M. V.; Musheev, M. U.; Krylov, S. N. Anal.Chem. 2009, 81, 490−494.(3) Ikebukuro, K.; Kiyohara, C.; Sode, K. Biosens. Bioelectron. 2005,20, 2168−2172.(4) Javaherian, S.; Musheev, M. U.; Kanoatov, M.; Berezovski, M. V.;Krylov, S. N. Nucleic Acids Res. 2009, 37, e62.(5) Ellington, A. D.; Szostak, J. W. Nature 1990, 346, 818−822.(6) Tuerk, C.; Gold, L. Science 1990, 249, 505−510.(7) Iliuk, A. B.; Hu, L.; Tao, W. A. Anal. Chem. 2011, 83, 4440−4452.(8) Hermann, T.; Patel, D. J. Science 2000, 287, 820−825.(9) Baker, B. R.; Lai, R. Y.; Wood, M. S.; Doctor, E. H.; Heeger, A. J.;Plaxco, K. W. J. Am. Chem. Soc. 2006, 128, 3138−3139.(10) Bi, S.; Zhou, H.; Zhang, S. Biosens. Bioelectron. 2009, 24, 2961−2966.

Analytical Chemistry Article

dx.doi.org/10.1021/ac300427d | Anal. Chem. 2012, 84, 5298−53035302

(11) Stojanovic, M. N.; de Prada, P.; Landry, D. W. J. Am. Chem. Soc.2000, 122, 11547−11548.(12) Xia, F.; Zuo, X.; Yang, R.; Xiao, Y.; Kang, D.; Vallee-Belisle, A.;Gong, X.; Yuen, J. D.; Hsu, B. B.; Heeger, A. J.; Plaxco, K. W. Proc.Natl. Acad. Sci. U.S.A. 2010, 107, 10837−10841.(13) Golub, E.; Pelossof, G.; Freeman, R.; Zhang, H.; Willner, I. Anal.Chem. 2009, 81, 9291−9298.(14) Tao, L.; Kennedy, R. T. Electrophoresis 1997, 18, 112−117.(15) Petrov, A.; Okhonin, V.; Berezovski, M.; Krylov, S. N. J. Am.Chem. Soc. 2005, 127, 17104−17110.(16) Drabovich, A. P.; Berezovski, M. V.; Musheev, M. U.; Krylov, S.N. Anal. Chem. 2009, 81, 490−494.(17) Connors, K. A. Binding Constants; John Wiley & Sons: NewYork, 1987.(18) Nguyen, T.-H.; Steinbock, L. J.; Butt, H.-; Helm, M.; Berger, R.d. J. Am. Chem. Soc. 2011, 133, 2025−2027.(19) Taylor, J. N.; Darugar, Q.; Kourentzi, K.; Willson, R. C.; Landes,C. F. Biochem. Biophys. Res. Commun. 2008, 373, 213−218.(20) Ashkin, A.; Dziedzic, J. M.; Bjorkholm, J. E.; Chu, S. Opt. Lett.1986, 11, 288.(21) Greenleaf, W. J.; Frieda, K. L.; Foster, D. A.; Woodside, M. T.;Block, S. M. Science 2008, 319, 630−633.(22) Baumann, C. G.; Smith, S. B.; Bloomfield, V. A.; Bustamante, C.Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 6185−6190.(23) Woodside, M. T.; Garcia-Garcia, C.; Block, S. M. Curr. Opin.Chem. Biol. 2008, 12, 640−646.(24) Liphardt, J.; Onoa, B.; Smith, S. B.; Tinoco, I., Jr.; Bustamante,C. Science 2001, 292, 733−737.(25) Neupane, K.; Yu, H.; Foster, D. A. N.; Wang, F.; Woodside, M.T. Nucleic Acids Res. 2011, 39, 7677−7687.(26) Anthony, P. C.; Perez, C. F.; Garcia-Garcia, C.; Block, S. M.Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 1485−1489.(27) Cecconi, C.; Shank, E. A.; Bustamante, C.; Marqusee, S. Science2005, 309, 2057−2060.(28) Koirala, D.; Dhakal, S.; Ashbridge, B.; Sannohe, Y.; Rodriguez,R.; Sugiyama, H.; Balasubramanian, S.; Mao, H. Nat. Chem. 2011, 3,782−787.(29) Koirala, D.; Yu, Z.; Dhakal, S.; Mao, H. J. Am. Chem. Soc. 2011,133, 9988−9991.(30) Yu, Z.; Schonhoft, J. D.; Dhakal, S.; Bajracharya, R.; Hegde, R.;Basu, S.; Mao, H. J. Am. Chem. Soc. 2009, 131, 1876−1882.(31) Luchette, P.; Abiy, N.; Mao, H. B. Sens. Actuators, B 2007, 128,154−160.(32) Smith, S. B.; Cui, Y. J.; Bustamante, C. Science 1996, 271, 795−799.(33) Mao, H.; Luchette, P. Sens. Actuators, B 2008, 129, 764−771.(34) Palassini, M.; Ritort, F. Phys. Rev. Lett. 2011, 107, 060601.(35) Dhakal, S.; Schonhoft, J. D.; Koirala, D.; Yu, Z.; Basu, S.; Mao,H. J. Am. Chem. Soc. 2010, 132, 8991−8997.(36) Jarzynski, C. Phys. Rev. Lett. 1997, 78, 2690−2693.(37) German, I.; Buchanan, D. D.; Kennedy, R. T. Anal. Chem. 1998,70, 4540−4545.(38) Berezovski, M.; Nutiu, R.; Li, Y.; Krylov, S. N. Anal. Chem. 2003,75, 1382−1386.(39) Huang, C.-C.; Cao, Z.; Chang, H.-T.; Tan, W. Anal. Chem.2004, 76, 6973−6981.(40) Szilagyi, A.; Bonn, G. K.; Guttman, A. J. Chromatogr., A 2007,1161, 15−21.(41) Krylov, S. N. Electrophoresis 2007, 28, 69−88.(42) Rundlett, K. L.; Armstrong, D. W. Electrophoresis 1997, 18,2194−2202.(43) Sassanfar, M.; Szostak, J. W. Nature 1993, 364, 550−553.(44) Junker, J. P.; Ziegler, F.; Rief, M. Science 2009, 323, 633−637.(45) Liphardt, J.; Dumont, S.; Smith, S. B.; Tinoco, I., Jr.;Bustamante, C. Science 2002, 296, 1832−1835.(46) Berezovski, M.; Krylov, S. N. J. Am. Chem. Soc. 2002, 124,13674−13675.(47) Clark, S. M.; Konermann, L. Anal. Chem. 2004, 76, 7077−7083.

(48) Stigler, J.; Ziegler, F.; Gieseke, A.; Gebhardt, J. C. M.; Rief, M.Science 2011, 334, 512−516.(49) Shank, E. A.; Cecconi, C.; Dill, J. W.; Marqusee, S.; Bustamante,C. Nat. Chem. 2010, 465, 637−640.(50) Lin, C. H.; Patel, D. J. Chem. Biol. 1997, 4, 817−832.

Analytical Chemistry Article

dx.doi.org/10.1021/ac300427d | Anal. Chem. 2012, 84, 5298−53035303